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The early development of vertebrate embryos is characterized by rapid cell proliferation necessary to support the embryo’s growth. During this period, the embryo must maintain a balance between ongoing cell proliferation and mechanisms that arrest or delay the cell cycle to repair oxidative damage and other genotoxic stresses. The ATM (Ataxia-Telangiectasia Mutated) kinase is a critical regulator of the response to DNA damage, acting through downstream effectors such as P53 and checkpoint kinases to mediate cell-cycle checkpoints in the presence of DNA damage. Mice and humans with inactivating mutations in ATM are viable, but have increased susceptibility to cancers. The possible role of ATM in limiting cell proliferation in early embryos has not been fully defined. One target of ATM and checkpoint kinases is the Cdc25 phosphatase, which facilitates cell cycle progression by removing inhibitory phosphates from cyclin-dependent kinases. We have identified a zebrafish mutant, standstill, with an inactivating mutation in cdc25a. Loss of cdc25a in the zebrafish leads to accumulation of cells in late G2 phase. We find that the novel family member, cdc25d, is essential for early development in the absence of cdc25a, establishing for the first time that cdc25d is active in vivo in zebrafish. Surprisingly, we find that cell cycle progression in cdc25a mutants can be rescued by chemical or genetic inhibition of ATM. Checkpoint activation in cdc25a mutants occurs despite the absence of increased DNA damage, highlighting the role of Cdc25 proteins to balance constitutive ATM activity during early embryonic development.
During the early phases of vertebrate embryo development, rapid cell proliferation is required to support the growth of the embryo and organogenesis. Throughout this time, the developing embryo is exposed to a variety of genotoxic stressors. External stressors include environmental toxins and chemicals as well as radiation, both ionizing and ultraviolet (1). Several endogenous sources of DNA damage have also been recognized, including oxidants, estrogens, alkylators and the products of lipid peroxidation and metabolic pathways (2, 3). Failure to repair the damage due to these extrinsic and intrinsic agents can lead to mutations that contribute to cancer, aging and degenerative diseases (1, 4).
In response to genotoxic stress, cells have evolved robust networks to detect DNA damage and to arrest or delay cell-cycle progression, allowing time for repair of the damage (5, 6). Testifying to the importance of these mechanisms, humans or mice with germline mutations in DNA damage response genes display a range of developmental defects, including growth retardation, microcephaly, cerebellar ataxia, sterility, bone marrow failure and immunodeficiency as well as predisposition to cancer (1, 4). Among the best characterized of these networks is the response to double-strand DNA (dsDNA) breaks, which is coordinated by the phosphoinositide-3-kinase-related protein kinase (PIKK) family member, ATM (Ataxia-telangiectasia mutated) (7). Upon sensing a dsDNA break or other change in chromatin structure, ATM phosphorylates key downstream targets including p53 and the checkpoint kinases CHK1 and CHK2, which in turn bring about cell-cycle arrest or delay to allow for repair through the Homologous Recombination or Non-Homologous End Joining pathways, or alternatively apoptosis and cell death (7–9). While ATM has clearly been shown to be important for the response to exogenous DNA damage, its possible roles in normal development are less clear. Similar to humans, mice deficient in ATM are viable but display growth retardation, neurologic dysfunction, infertility, defective T lymphocyte maturation, radiation sensitivity and increased cancer incidence (10, 11). Exposure to teratogens such as phenytoin or genotoxic insults such as gamma radiation impairs the development of ATM-null embryos, and simultaneous deletion of ATM and other DNA damage repair factors including histone H2AX and DNA-dependent protein kinase is embryonic lethal in mice (12, 13). However, owing to the relative inaccessibility of mammalian embryonic development, it is unclear what roles ATM may play during the course of normal embryogenesis. In particular, it is not known to what degree ATM-dependent checkpoint mechanisms may oppose cell proliferation during early embryogenesis.
One of the key targets of the ATM-CHK1/2 network is the CDC25 family of phosphatases, key regulators of the cell cycle. Under conditions that favor cell-cycle progression, CDC25 removes inhibitory phosphates from cyclin-dependent kinases (CDKs), allowing cyclin/CDK complexes to drive cell-cycle transitions (14). In response to dsDNA breaks, CHK1 and CHK2 phosphorylate CDC25, targeting it for destruction and impeding CDK activation (15). Mammalian cells express three different CDC25 genes: CDC25A, CDC25B and CDC25C. CDC25A appears to act alone in controlling entry from G1 to S and intra-S progression, while all three CDC25 genes function in G2/M progression (16).
The developmental roles of Cdc25 have been partially defined. In mice, Cdc25A is essential for embryonic development. Cdc25A−/− embryos are resorbed at around E6.5 due to widespread apoptosis (17). Although Cdc25A−/− mice die early in development, mice lacking Cdc25B and Cdc25C survive with normal cell cycle progression and checkpoint function (18). Therefore, Cdc25A likely compensates for the other Cdc25 genes and may be the most functionally important mammalian Cdc25. Unlike mammals, zebrafish express a single canonical CDC25, designated cdc25a, during embryogenesis (19). In zebrafish embryos, overexpression of cdc25a drives cells into mitosis (19) and blocks cell cycle lengthening and acquisition of G2 phase as early embryonic cell cycles give rise to post-midblastula transition, asynchronous cell cycles (20). Zebrafish also express a divergent family member, designated cdc25d, which is homologous to cdc25a but is not present in mammals (19). cdc25d can rescue a fission yeast cdc25ts mutant, but has not been shown to have detectable activity in zebrafish (19, 20). It is not known whether the single canonical zebrafish cdc25a is essential for development, or whether cdc25a and cdc25d have any redundant roles in cell-cycle regulation, nor is it known if zebrafish cdc25 family members participate in DNA damage checkpoints.
Previously, we performed a screen for mutations that affect embryonic cell proliferation in zebrafish (21). Here, we report the identification of an inactivating mutation in zebrafish cdc25a. cdc25a is essential for zebrafish embryonic development, and cdc25a-deficient mutants accumulate cells in G2/M phase. The cell-cycle defects of cdc25a-deficient mutants can be partially rescued by cdc25d, demonstrating in vivo activity of this divergent family member. We reasoned that this model could help us to understand epistatic relationships of CDC25 to other checkpoint genes in a whole organism that is not subject to extrinsic genotoxic stress. We find that chemical or genetic inhibition of ATM rescues the accumulation of cells in G2/M phase in cdc25a-deficient embryos. ATM is activated in the cdc25a mutants despite the absence of widespread DNA double-strand breaks, and we present evidence that ATM also impedes cell-cycle progression in embryos with wildtype levels of CDC25 activity. These results emphasize the important balance between mechanisms that favor cell proliferation and the ATM-mediated checkpoint response during early embryonic development in vertebrates.
Zebrafish were maintained according to standard procedures (22). All work with zebrafish was carried out under protocols approved by the Institutional Animal Care and Use Committees at UT Southwestern Medical Center, an AAALAC-accredited institution.
24-hour old embryos were dechorionated, euthanized with tricaine and fixed in 4% paraformaldehyde (PFA) in 1× phosphate-buffered saline (PBS) overnight at 4°C. Immunohistochemistry was performed using 1:1000 anti-phosphohistone H3 Serine 10 (pH3) (Santa Cruz Biotechnology, Santa Cruz, CA, catalog no. sc-8656-R); 1:200 anti-Mouse Cdc25A (Santa Cruz Biotechnology catalog no. sc-97) or 1:1000 zebrafish-specific anti-phosphohistone H2AX (23) followed by incubation with 1:350 Horseradish peroxidase conjugated goat anti-rabbit IgG (Jackson Immunochemicals, Jackson ME) and staining with diaminobenzidine (DAKO, Carpinteria, CA) for 10 minutes according to the manufacturer’s protocol. For fluorescent imaging, secondary antibody was a 1:15,000 dilution of Alexafluor-488 conjugated goat anti-rabbit IgG (Invitrogen, Carlsbad, CA). TUNEL assay was performed using the Apoptag Red In Situ Apoptosis Detection Kit (Millipore, Billerica, MA) as described (24). Acridine orange staining was performed as described (24). Immunoblots were performed with rabbit anti-phospho-CHK1 (Ser435) from Cell Signaling Technology (Danvers, MA). The antibody recognizes a protein of 50 kD in lysates of zebrafish embryos, in good agreement with the predicted size of zebrafish chk1 (410 amino acids; Accession: NP_956487.1), and was previously used to detect activated chk1 protein in lysates of zebrafish tissue (25). Anti-phospho-Rb (Ser 795) is validated for recognition of phosphorylated zebrafish Rb protein (www.cellsignal.com/products/9301.html) and was previously shown to recognize phospho-Rb in zebrafish (26)
24-hour or 48-hour zebrafish embryos were disaggregated to a single-cell suspension as described (27). Zebrafish cells were embedded in low-melt agarose gels, lysed and then electrophoresed using the Trevigen CometAssay kit according to the manufacturer’s protocol (Trevigen, Gaithersburg, MD).
12-hour old embryos were treated with 10µM ATM kinase inhibitor KU55933 (Calbiochem, San Diego, CA) in E3 zebrafish embryo medium (22) and 1% DMSO for 12 hours at 28°C, then euthanized with tricaine, fixed in 4% PFA/1× PBS overnight at 4°C and processed for immunohistochemistry. To quantify anti-pH3 staining, 12 images were taken for each treatment and genotype. The pH3-positive cells in an area extending from the end of the yolk tube extension to the tip of the tail were independently counted for each embryo by two observers who were blinded to the identity of the samples. All counts were normalized by the total area (measured using ImageJ software) to account for differing size of the tail regions.
The human non small-cell lung cancer cell line A549 (kindly provided by Dr. J. Minna) was maintained in RPMI media supplemented with 10% Fetal Bovine Serum. For overexpression of Cdc25, cells were transfected with pCMV-SPORT6 containing a full-length Mus musculus Cdc25A cDNA (Accession number BC046296, Open Biosystems, Huntsville, AL). Immunoblotting was performed using 1:1000 pS1981-ATM (Rockland, Gilbertsville, PA) or 1:500 anti-gH2ax (Millipore, Billerica, MA) as described (28).
0.5mM Cdc25a 5’UTR (TAATCAGCCAGGCGCGATTAAGAAC), cdc25a splice site (ATGACAACCTCACCTCAGCCATGTT), control (CCTCTTACCTCAGTTACAATTTATA) or cdc25d 5’UTR (AATCTCCAGCGCATCACCGGCCATT) morpholinos were injected into 1–2 cell zebrafish embryos from the AB strain or from offspring of cdc25a+/− adults. Morpholinos were purchased from Gene-Tools (Philomath, OR). ATM, CDKN1A and ATR morpholinos were as described (23, 29). The genotypes of injected embryos were confirmed by PCR using primers flanking the cdc25a mutation site (GGTGTTTGACTCCAATCTGCT and CAACAAGCACAGGCTAATGG) followed by digestion of the PCR product with BsiEI (New England Biolabs, Ipswich, MA) and gel electrophoresis.
Previously, we described a forward-genetic screen to identify mutations that altered embryonic cell proliferation in the haploid F2 progeny of ENU-mutagenized zebrafish (21, 30). Using whole-mount immunohistochemistry with an antibody specific for the phosphorylated form of histone H3 (to evaluate cell proliferation), seven mutant lines were identified including slycz61, sfdcz213, mybl2cz226, espl1cz280, llgcz3322, sdscz319 and slhcz333 (21). The standstill mutant (sdscz319) was identified based on a markedly decreased fraction of phosphohistone H3 (pH3)-positive cells in the mutant embryos (Figure 1). The standstill mutant morphologic phenotype becomes evident at approximately 24 hours post-fertilization (hpf) and includes microcephaly, microphthalmia, abnormal body shape and reduced pigmentation (Supplementary Figure S1). To further understand the cell proliferation defect, we performed flow cytometric DNA content analysis, which demonstrated an accumulation of cells in the G2/M phase of the cell cycle (Figure 1E). Histone H3 is phosphorylated in late G2 phase, coincident with the onset of chromatin condensation and remains phosphorylated until the chromosomes begin to decondense in anaphase (31, 32). Thus, the standstill mutation appears to result in accumulation of cells in the G2 phase of the cell cycle, prior to the onset of chromatin condensation (33).
To understand the molecular nature of the defect in standstill embryos, we used meiotic recombinational mapping to identify the mutant locus. We performed bulk segregant analysis (34) followed by intermediate-resolution mapping with a panel of 88 mutant embryos to assign standstill to zebrafish chromosome 13. For high-resolution mapping, we assembled a panel of 1,758 phenotypically mutant embryos and localized the mutation to an approximately 5 cM interval between microsatellite markers Z6259 and ZJA5 (Figure 2). The map position was further refined with a combination of known markers and novel microsatellites derived from genomic sequence. The critical interval contains a single gene, the zebrafish ortholog of Cdc25A (19). We sequenced the cdc25a gene from wildtype and standstill mutant embryos and found that the mutants contain a C to T mutation at codon 206 of the coding sequence, which is predicted to result in a premature stop codon and a truncated protein lacking the catalytic domain (Figure 2B,C). standstill mutant embryos did not stain with an anti-mouse Cdc25A antibody (Figure 2E), and whole-mount in situ hybridization revealed reduced cdc25a mRNA in standstill mutants, consistent with nonsense-mediated decay (Supplementary Figure S2)
To confirm that cdc25a deficiency was responsible for the standstill phenotype, we designed a morpholino oligonucleotide (MO) targeting the splice junction between exon 1 and intron 1 of the cdc25a pre-mRNA. cdc25a MO-injected embryos displayed a phenotype identical to the standstill mutant, including microcephaly, microphthalmia and bent tail. Immunohistochemical staining for pH3 showed that the MO-injected embryos had reduced cell proliferation, phenocopying the standstill mutation (Figure 2G). We obtained identical results with a translation-blocking MO directed against the cdc25a initiating ATG (Supplementary Figure S3). Thus, standstill is encoded by zebrafish cdc25a.
During the cell cycle, Cdc25 removes inhibitory phosphates from cyclin-dependent kinases to allow cell-cycle progression. Despite lacking a key cell cycle regulator, cdc25a−/−-deficient embryos undergo gastrulation and relatively normal organogenesis. The ability of embryos to complete early stages of development could be due in part to residual maternally-supplied wild type cdc25a. However, early development also occurs in embryos injected with translation-blocking MO (Supplementary Figure S3), which is predicted to inhibit translation of both maternal and zygotic mRNAs. We investigated whether cdc25a deficiency is partially compensated by activity of cdc25d, the only other identified zebrafish cdc25. cdc25d has been characterized as a highly divergent isoform of cdc25. Zebrafish cdc25d functionally complements Schizosaccharomyces pombe cdc25 activity (19), however its function in zebrafish has not been established. MO knockdown of cdc25d alone led to a mild growth defect (Figure 3C), but knockdown of cdc25d in cdc25a−/− embryos led to a much more significant growth defect (Figure 3D), as did double knockdown of cdc25a and cdc25d in wildtype embryos (Figure 3E). These results indicated that cdc25d activity contributes to the early development of cdc25a−/− embryos. Based on these results, we asked whether overexpression of cdc25d might at least partially rescue the cell proliferation defect of cdc25a−/− embryos. To test this possibility, we injected cdc25a−/− embryos with cdc25d mRNA and stained for pH3 (Figure 3F,G). Embryos injected with cdc25d mRNA had a slightly increased number of pH3 foci compared to control cdc25a−/− embryos (the number of pH3-positive cells in a defined area of the tail was 19.3±11.3 for cdc25a−/− embryos and 35.1±13.1 for cdc25a−/− embryos injected with cdc25d mRNA; p=0.059 by two-tailed t-test). cdc25d mRNA did not increase the number of pH3-positive cells in wt embryos (118.5±22.3 pH3-positive cells in control vs. 131±19.6 in cdc25d mRNA injected; p=0.39). cdc25d mRNA injection did not rescue morphology or survival in cdc25a−/− embryos. Some caution is warranted in interpreting the cdc25d gain of function experiments, because they relied on overexpression and because the rescue was incomplete. However, taken together with the loss of function data, these results strongly suggest that cdc25 activity is necessary for early embryonic development in zebrafish, and that cdc25a and cdc25d are at least partially redundant.
While overexpression of cdc25d increased the number of mitotic cells in cdc25a−/− mutants, endogenous levels of cdc25d activity were unable to rescue cdc25a deficiency, despite the fact that cdc25d is expressed during early development (19). It is possible that cdc25d activity is simply too weak to complement cdc25a deficiency. However, we also considered the possibility that another factor inhibits cell proliferation in cdc25a mutants. We noted that the phenotype of cdc25a−/− mutant embryos (accumulation in G2 phase prior to the onset of chromatin condensation as signaled by pH3 positivity) is also compatible with cell-cycle arrest due to activation of the G2/M checkpoint. The G2/M checkpoint is dependent on the action of the PI3-kinase family member ATM; it is present in zebrafish embryos and is robustly activated by DNA-damaging agents such as gamma radiation (Supplementary Figure S4).
To test whether the G2/M checkpoint is activated in cdc25a−/− embryos, we treated the embryos with the ATM inhibitor KU55933. To demonstrate that KU55933 inhibits zebrafish ATM, we pretreated wildtype embryos with 10µM KU55933 for four hours, then exposed pretreated embryos to 12 Gy ionizing radiation.. At 48 hpf, irradiated KU55933-treated embryos displayed severe morphologic defects, similar to those present in irradiated embryos after MO knockdown of ATM (29) (Supplementary Figure S5).
Having established that Ku55933 is active in fish, we next tested whether the G2/M accumulation in cdc25a−/− mutants was the result of activation of the G2/M checkpoint. We crossed heterozygous cdc25a+/− fish and treated the resulting embryos with KU55933 from 12 hpf to 24 hpf, then fixed the embryos and stained for pH3 (Figure 4). pH3 levels were quantified by counting the number of pH3 foci in a defined area of the embryo (Figure 4 G). Control DMSO-treated clutches displayed the expected 25% of embryos with a very low number of pH3-positive cells as compared to wild type embryos (Figure 4A,C,G). In contrast, cdc25a−/− embryos treated with KU55933 exhibited a marked increase in the number of pH3-positive cells (Figure 4D,G) (p< 0.0001). We obtained identical results by inhibiting ATM with caffeine (Supplementary Figure S6). We confirmed that the embryos with increased cell proliferation after KU5593 treatment were genotypically cdc25a−/− using a Restriction Fragment Length Polymorphism specific for the standstill cdc25a mutant allele (not shown).
To rule out an off-target effect of KU55933 and provide an independent means of inhibiting ATM, we injected 0.2mM control or ATM-specific MOs (29) into offspring of heterozygous cdc25a+/− fish, and again assessed cell proliferation using pH3. Compared to control MO injected embryos, ATM MO injected embryos had increased cell proliferation (Figure 4E,F). Further confirming these results, we found that cdc25a−/− embryos contain elevated levels of phospho-Y15-Cdc2, and that inhibition of ATM results in reduction of this inhibitory phosphorylation (Figure 4H). We also considered the possibility that the phenotype of cdc25a−/− mutants is due at least in part to activation of a replication checkpoint due to replication stress. To test this possibility, we knocked down expression of atr (Ataxia-telangiectasia and rad9-related), another PI3K family member that delays or arrests the cell cycle in response to stalled replication forks or other forms of replication stress (25). Knockdown of atr did not rescue the proliferation defect of cdc25a morphants (Figure 4G). Furthermore, cdc25a morphants did not exhibit elevated levels of the phosphorylated form of CHK1 protein, as would be expected if the replication checkpoint were activated (Supplementary Figure S7E).
To test whether the increased proliferation in KU55933-treated cdc25a−/− embryos was due to a true increase in cell cycle progression or merely indicative of a release of the G2/M checkpoint into mitosis, we performed FACS analysis on cdc25a−/− embryos and phenotypically wildtype clutchmates treated either with the KU55933 or 1% DMSO (samples 1 and 2, Figure 4J). While cdc25a−/− embryos treated with DMSO accumulated in G2 (sample 3), cdc25a−/− embryos treated with the ATM inhibitor (sample 4) exhibited a large G1 peak, indicating they had resumed the cell cycle. Quantification of the FACS plots revealed the following distributions: cdc25a−/− in DMSO: 30.6%G1, 17.8% S, 64.6% G2; cdc25a−/− in KU55933: 61.5%G1, 31.2%S, 12.8%G2. For comparison, the values for wildtype in DMSO are: 49.7%G1, 32.2% S, 17.6% G2; and wildtype in KU55933: 49.7%G1, 36.1% S, 15.8% G2. The large fraction of cells in G1 in KU55933-treated cdc25a−/− embryos likely reflects the release of a large number of G2 cells into mitosis and a subsequent G1.
These results indicated that G2/M accumulation in cdc25a−/− embryos was largely due to activation of the ATM-dependent G2/M checkpoint, and that cell proliferation in the mutant embryos could resume once ATM activity was inhibited. Having demonstrated that cdc25a and cdc25d activities are at least partially redundant (Figure 3), we hypothesized that in KU55933-treated, cdc25a-deficient embryos, cdc25d activity drives the G2/M transition. To test this model, we knocked down cdc25d in cdc25a−/− embryos and tested whether the cell cycle resumed after ATM inhibition. We found that, in the absence of expression of either cdc25a or cdc25d, KU55933 treatment failed to rescue the cell proliferation defect (Figure 4G,J,K). Thus, in the absence of cdc25a activity, ATM appears to exert its regulatory effect on cdc25d.
cdc25 activity is also regulated at the G1/S checkpoint, principally by p53 and its downstream target p21 (8). To determine whether activation of the G1/S checkpoint contributed to the cell-cycle phenotype of cdc25a-deficient embryos, we knocked down cdc25a in p53M214K mutants, which are deficient in p53-mediated responses (35). We also simultaneously knocked down cdc25a and cdkn1a, which encodes p21. Neither p53 deficiency nor p21 deficiency increases the number of proliferating cells in cdc25a-deficient embryos (Supplementary Figure S7A,B). Finally, we stained wildtype and cdc25a−/− embryos for the phosphorylated form of Retinoblastoma protein, an indicator of G1/S cyclin-Cdk activity. Rb protein was phosphorylated in both wildtype cdc25a-deficient embryos, indicating that the G1/S transition was occurring in the mutants (Supplementary Figure S7C,D).
Having established that an ATM-dependent G2/M checkpoint is present in cdc25a−/− embryos, we investigated the mechanism of checkpoint activation. We first considered the possibility that Cdc25a deficiency results in increased DNA double strand breaks in developing embryos. We employed the comet assay to measure the amount of DNA damage on a single-cell basis (36). We have recently adapted this method to measure DNA damage in zebrafish cells (24). Performing the comet assay on 24hpf cdc25a−/− embryos, wildtype embryos or wildtype embryos treated with 12 Gy of ionizing radiation as a positive control, we observed a slight decrease in spontaneous DNA damage in the cdc25a−/− mutants (Figure 5A–D). While it is possible that the comet assay fails to detect minor amounts of DNA damage, these data indicate that loss of cdc25a itself does not induce severe DNA damage. The observed decrease in comet tail size in cdc25a−/− mutants may be due to decreased spontaneous replication-related DNA damage due to G2/M cell cycle arrest.
As an alternative approach to detect DNA damage foci in zebrafish, we developed an anti-phospho-Histone H2AX rabbit polyclonal antibody specific for phosphorylated zebrafish Histone H2AX (23). Phosphorylated Histone H2AX, designated γH2AX, is present at histone cores flanking double-stranded breaks (37). We performed immunoblot analysis on wild type and cdc25a−/− embryos to measure γH2AX levels (Figure 5E). Compared to wild type controls, cdc25a−/− embryos have similar to decreased γH2AX levels. As expected, irradiated wild type embryos exhibit increased γH2AX, which can be reduced by pre-treatment of the embryos with KU55933. Therefore, the ATM-γH2AX axis is present and normally functioning in zebrafish, but cdc25a−/− embryos do not exhibit increased DNA damage, as measured by Histone H2AX phosphorylation. The low level of γH2AX in cdc25a−/− embryos made it unlikely that widespread DNA damage could account for the observed profound cell cycle arrest. As further evidence of this point, we tested whether H2AX phosphorylation precedes the onset of the cell cycle phenotype in cdc25a−/−, by staining age-matched embryos for γH2AX and pH3 (Supplementary figure S8). pH3 levels are detectably lower at 90% epiboly stage (about 8 hpf) and continue to progressively decrease. In contrast, H2AX phosphorylation does not become detectable until the 14 somite stage. These data indicate that cell cycle arrest precedes the an increase in apoptotic γH2AX.
The above results indicate that cdc25a deficiency triggers an ATM-dependent G2/M accumulation in developing embryos, but the accumulation is not attributable to increased DNA double-strand breaks. To account for ATM activity in cdc25a−/− embryos in the absence of detectable DNA damage, we next considered the possibility that Cdc25A directly regulates ATM via Cdc25’s phosphatase activity. To test the possibility that Cdc25A directly dephosphorylates and regulates ATM, we overexpressed mouse Cdc25A in A549 cells for 48h, irradiated the cells and assessed the phosphorylation status of ATM and its target, CHK2, by immunoblotting (Supplementary Figure S9). Both ATM and CHK2 demonstrated increased phosphorylation status after irradiation, and overexpression of Cdc25A did not decrease the phosphorylation, indicating that Cdc25A does not directly regulate ATM. We also noted increased phospho-ATM in non-irradiated Cdc25A-overexpressing cells consistent with previous findings that overexpression of Cdc25A leads to increased DNA damage (38).
Finally, we investigated whether ATM is generally active during early developmental cell cycles, and if its effects on the cell cycle were being unmasked by cdc25a deficiency in the mutant embryos. Indeed, wildtype embryos treated with KU55933 and quantified for the number of mitotic cells using pH3 as a marker exhibited an increased number of mitotic cells compared to controls (Figure 4A,B,G), however this effect did not reach statistical significance (p=0.09). These results suggest that, during normal development, ATM is active and may attenuate the percentage of cells progressing from G2 to M phase. Based on the above results, we conclude that ATM is constitutively active during development, but does not completely deplete Cdc25 activity. Thus, in wildtype animals a balance between cell cycle arrest/delay and cell cycle progression is maintained. cdc25a deficiency unmasks ATM activity, leading to G2 accumulation and impaired embryonic development.
In performing this analysis, we noted that embryos treated with 10µM KU55933 demonstrated some developmental toxicity in the absence of irradiation, suggesting ATM function is necessary for healthy development of zebrafish. To explore this possibility further, we knocked down ATM expression with a morpholino and monitored developmental toxicity and survival in the injected embryos (Supplementary Figure S10). Compared to control MO-injected embryos, ATM morphants displayed increased developmental defects and significantly lower survival. This result is consistent with an earlier report of lower survival in ATM morphants (29) and suggests ATM may play an important role in normal development, even in the absence of exogenous genotoxic stress.
ATM was first identified as the protein mutated in the clinical syndrome Ataxia-Telangiectasia, which is characterized by immune dysfunction, abnormal sensitivity to ionizing radiation, cerebellar ataxia and telangiectases. Subsequent work revealed that ATM and the related phosphatidyl kinase kinases (PIKKs), ATR and DNA-PK, are central regulators of the DNA damage response (39, 40). In response to genotoxic stresses causing DNA double-strand breaks, one action of ATM is to mediate G2/M cell-cycle arrest by acting through the checkpoint kinases CHK1/CHK2 to target Cdc25 for destruction (41–43). In keeping with this role of ATM in maintaining genomic integrity, people with mutations in the ATM gene are at increased risk of developing cancer (44, 45).
More recently, it has become clear that the PIKKs and their downstream partners have important roles in maintaining normal cell homeostasis, even in the absence of exogenous sources of DNA damage. ATM and ATR control the timing of replication origin firing, in the absence of DNA damage (46). CHK1 localized to the centrosome has been shown to regulate Cdc25B activity to control the entry into mitosis (47). Constitutive Histone H2AX phosphorylation and constitutive ATM activation (CAA) in normal cycling cells has been postulated to occur, and has been attributed in interphase cells to DNA damage due to endogenous oxidative stress (48–50). In mitotic cells, CAA and H2AX phosphorylation have been associated with chromatin condensation, and postulated to play a role in maintaining mitotic spindle integrity (51, 52).
Here, we show that a G2/M accumulation is present in developing cdc25a mutant embryos, as evidenced by loss of pH3 positive nuclei and by DNA content profiling. This accumulation is due to activation of ATM and can be partially overcome by inhibiting ATM, so long as cdc25d activity is present. The G2/M accumulation is not due to increased DNA damage, but appears to be constitutive, as inhibiting ATM in wildtype embryos also increases the number cells making the G2-to-M transition. In the cdc25a−/− mutants, we do not believe that ATM activation results from premature chromatin condensation or other mitotic abnormalities, because cells in developing mutant embryos accumulate prior to the onset of chromatin condensation as evidenced by markedly reduced levels of Histone H3 Serine 10 phosphorylation.
We hypothesize that a basal level of ATM activity occurs in normal cell cycles. In cdc25a−/− cells, early embryonic development proceeds fairly normally due to compensatory phosphatase activity of cdc25d. In cells with normal levels of cdc25 activity, the basal level of ATM activity subtly attenuates cell cycle progression, however, in cdc25a−/− embryos, the phosphatase activity of cdc25d is overwhelmed by basal ATM activity, leading to cell cycle accumulation in the absence of marked DNA damage (Figure 6). Only when ATM activity is genetically or pharmacologically abrogated can cell cycle activity resume. Thus, we propose that a hierarchy of Cdc25 activity is present, such that enough combined cdc25 phosphatase activity must be present to overcome the basal ATM activity to progress the cell from G2 to M phase. Our data do not indicate the mechanism(s) regulating cdc25d. The sequence of cdc25d is poorly-conserved relative to other Cdc25 family members (19), and we did not identify conserved consensus phosphorylation sites.
Because of our observation that G2/M accumulation is likely due to basal ATM activity, we sought the mechanism by which Cdc2 is being de-phosphorylated in ATM-inhibited embryos. We discovered that zebrafish cdc25d is expressed and is able to mediate cell cycle progression to a modest degree. Knockdown of cdc25d had profound consequences for development in cdc25a−/− or cdc25a knockdown embryos, the first demonstration that cdc25d is active in vivo in zebrafish. Ectopic expression of cdc25d led to modest increases in the number of proliferating cells that was detectable in cdc25a−/− embryos. Previous reports indicated that cdc25d could rescue a yeast cdc25 mutant but failed to show activity in zebrafish (19, 20). Consistent with these results, we did not detect a significant effect in wt embryos, suggesting that cdc25a is responsible for the majority of Cdc25 activity during development. Expression of cdc25d did not, however, rescue the morphologic phenotype of cdc25a−/− embryos. This finding may be due to limited ability of cdc25d to oppose ATM activity in cdc25a−/− mutants, or may point to essential roles of cdc25a in processes other than G2/M progression.
The early development of vertebrate embryos requires rapid and highly regulated cell divisions. Cdc25 phosphatases are important mediators of this rapid cell-cycle flux. Balancing this rapid growth is the need to respond to genotoxic insults from exogenous or endogenous, to ensure the health and genomic stability of the developing embryo. The results presented here emphasize the delicate balance between cell proliferation and cell cycle arrest/delay, and highlight the competing roles of ATM and Cdc25 family members in this process. The cdc25a−/− mutant also provides a sensitive model of physiologic ATM activity that could be used to further dissect the genetics of the DNA damage response and to screen for novel checkpoint inhibitors.
We thank Reinhard Kodym for helpful discussions and advice, Nuno Gomez for assistance with the comet assay, Elizabeth Patton for comments on the manuscript and Leonard Zon for generously supporting the early phase of this work. DV was supported by NIH training grant 5 T32 GM08203. This work was supported by the Amon G. Carter Foundation and by grant 1R01CA135731 from the National Cancer Institute to JFA.
The abbreviations used are:
The authors declare no Conflict of Interest